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© 2006 The Authors
Fatty acid metabolism in
adipose tissue, muscle and
liver in health and disease
Keith N. Frayn*1, Peter Arner† and
Hannele Yki-Järvinen‡
*Oxford Centre for Diabetes, Endocrinology and Metabolism,
University of Oxford, Churchill Hospital, Oxford OX3 7LJ, U.K.,
†Department of Medicine, Karolinska Institute, Huddinge University
Hospital, S 141 86 Huddinge, Sweden, and ‡Department of Medicine,
Division of Diabetes, University of Helsinki, PO Box 340, 00029
Helsinki, Finland
Abstract
Fat is the largest energy reserve in mammals. Most tissues are involved in
fatty acid metabolism, but three are quantitatively more important than
others: adipose tissue, skeletal muscle and liver. Each of these tissues has a
store of triacylglycerol that can be hydrolysed (mobilized) in a regulated
way to release fatty acids. In the case of adipose tissue, these fatty acids
may be released into the circulation for delivery to other tissues, whereas in
muscle they are a substrate for oxidation and in liver they are a substrate for
re-esterification within the endoplasmic reticulum to make triacylglycerol
that will be secreted as very-low-density lipoprotein. These pathways are
regulated, most clearly in the case of adipose tissue. Adipose tissue fat storage
is stimulated, and fat mobilization suppressed, by insulin, leading to a drive to
store energy in the fed state. Muscle fatty acid metabolism is more sensitive
to physical activity, during which fatty acid utilization from extracellular and
1To
whom correspondence should be addressed (email [email protected]).
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intracellular sources may increase enormously. The uptake of fat by the liver
seems to depend mainly upon delivery in the plasma, but the secretion of
very-low-density lipoprotein triacylglycerol is suppressed by insulin. There is
clearly cooperation amongst the tissues, so that, for instance, adipose tissue fat
mobilization increases to meet the demands of skeletal muscle during exercise.
When triacylglycerol accumulates excessively in skeletal muscle and liver,
sometimes called ectopic fat deposition, then the condition of insulin resistance
arises. This may reflect a lack of exercise and an excess of fat intake.
Introduction
Triacylglycerol (or triglyceride) is a concentrated form in which to store
energy. Triacylglycerol circulates in the plasma in lipoproteins and is also
present as lipid droplets within cells. The ‘energy density’ of lipid stored
in cells in this way is approx. 37 kJ/g. The energy density of whole adipose
tissue (with cytoplasm, connective tissues etc.) is approx. 30 kJ/g. In contrast,
glycogen in pure form has an energy density of approx. 17 kJ/g, but when
stored in hydrated form with about three times its own weight of water (as
it is in cells), this reduces to approx. 4 kJ/g. There are three main organs that
store triacylglycerol in a regulated way and hydrolyse it to release fatty acids,
either for export or for internal consumption. These are, in order of amount
of triacylglycerol typically stored, adipose tissue, skeletal muscle and the
liver. Recent years have seen an enormous increase in our understanding of
Dietary fat
Muscle
Small intestine
Ox
LPL
Chylomicrons
LPL
FA
TG
Chylomicrons
VLDL
NEFA
Remnants
Remnants
LPL
?DNL
DNL
NEFA
TG
FA
Ox
Cytosolic
ER-TG
TG
LPL
VLDL
Liver
Adipose tissue
Figure 1. Lipid exchanges between gut, adipose tissue, skeletal muscle and liver
Note that adipose tissue LPL also releases NEFA into the plasma, making further fatty acids available for uptake by muscle and liver. ER, endoplasmic reticulum; FA, fatty acids; Ox, ␤-oxidation.
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Table 1. Characteristics of fatty acid and TG metabolism compared in
adipose tissue, skeletal muscle and liver
Adipose tissue
Skeletal muscle
Liver
(at rest)
Input
LP–TG ~ 45 g/day
NEFA ~ 20 g/day
NEFA ~ 5 g/day
LP–TG ~ 10 g/day
NEFA ~ 20 g/day
Remnant-LP–TG
~ 25 g/day
DNL ~ 1 g/day
Stimulation of uptake
Feeding/insulin
Fasting (high NEFA
Delivery
supply)
Exercise
FA transporters
FAT
FAT
FAT
FATP-1, 4
FATP-1, 4
FATP-2, 5
15 kg
300 g
100 g
Half-life of store
250 days
24 h
100 h
Lipolysis of TG store
HSL - stimulated lipolysis HSL ?
Typical whole-body TG
store
Unknown
ATGL - basal lipolysis
MGL - monoacylglycerol
hydrolysis
Releases mainly
NEFA into plasma
FA for oxidation
FAs re-esterified in
ER and released as
VLDL-TG
Quantitative data involve some estimates and should be taken as representative only. ER,
endoplasmic reticulum; FA, fatty acids; LP, lipoprotein; MGL, monoacylglycerol lipase; TG,
triacylglycerol.
triacylglycerol and fatty acid metabolism in these organs, and of its relationship
to health and disease.
There is a continual flow of fatty acids between the three tissues described
above, outlined in Figure 1 with some quantitative estimates in Table 1. Dietary
fat enters the plasma from the small intestine in the form of chylomicron-triacylglycerol. The chylomicrons are the largest of the lipoprotein particles and
have a very fast turnover (the half-life of chylomicron-triacylglycerol in the
circulation is around 5 min). They deliver triacylglycerol-fatty acids to tissues expressing the enzyme LPL (lipoprotein lipase), an enzyme bound to
the endothelial cells lining capillaries of a number of tissues including adipose tissue, skeletal and cardiac muscle (more details are given in Figure 2).
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Capillary
Lipoprotein
lipase
Endothelial cells
Secretory
vesicle
Nucleus RER
Adipocytes
mRNA
Golgi
Fat droplet
Chylomicron
Capillary
TG
TG
VLDL
Fatty acids
TG
Lipoprotein
lipase
Endothelial cells
TG
Adipocytes
Figure 2. Regulation and action of LPL
This is shown for adipose tissue but aspects of the regulation and action are likely to be similar
in other tissues. Top panel: LPL is synthesized in the rough endoplasmic reticulum (RER) and
post-translationally modified, becoming active in the Golgi apparatus. An important aspect of
activation is dimerization. From the Golgi, LPL is secreted via the vascular endothelial cells to
proteoglycan chains attached to the endothelial cell membrane. The lipid droplet of the mature
adipocytes has been reduced in size in this diagram for clarity. In principle, regulation of LPL
activity could occur at many points. In fact, up-regulation of adipose tissue LPL activity in the
fed state occurs with little change in mRNA or intracellular enzyme mass. It appears mainly to
involve the mobilization of active enzyme to the capillary-luminal aspect of the endothelial cells.
There is no evidence for reversible phosphorylation as a means of regulation of LPL activity. In
fasting, more LPL becomes inactive (perhaps via monomerization). Note that, in contrast, LPL in
skeletal and cardiac muscle is activated during fasting. Lower panel: LPL acts via binding to the
triacylglycerol-rich lipoprotein particles, chylomicrons (carrying dietary fat) and VLDL (secreted
from the liver). It hydrolyses the triacylglycerol to liberate fatty acids, that can be taken up by
the adipocyte (or muscle cell) and esterified to form new triacylglycerol. In skeletal or cardiac
muscle, these fatty acids might also be a substrate for oxidation. Lower panel taken from Frayn,
K.N. (2003) Metabolic Regulation : a Human Perspective, 2nd edn. Published by Blackwell Publishing.
The remaining ‘remnant’ particles (that have lost perhaps two-thirds of their
triacylglycerol) are mainly taken up by the liver. This is the starting point for
fatty acid metabolism in the tissues.
Adipose tissue takes up fatty acids via the action of LPL. It releases fatty
acids by the hydrolysis (lipolysis) of stored (intracellular) triacylglycerol, by
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the action of the enzyme known as HSL (hormone-sensitive lipase) and other
lipases, to liberate NEFA (non-esterified fatty acids) into the plasma. Plasma
NEFA circulate bound to albumin. The turnover of plasma NEFA is also
rapid, with a half-life of 4–5 min. The liver takes up fatty acids in various forms
and liberates triacylglycerol-fatty acids as VLDL (very-low-density lipoprotein) particles. In this scenario, skeletal and heart muscle (and other oxidative
tissues not considered here) are the only pure consumers, taking up fatty acids
either from lipoprotein-triacylglycerol or from the plasma NEFA pool and
using them ultimately for oxidation.
Regulation of triacylglycerol and fatty acid metabolism at rest
Overview
The pattern of flow of fatty acids between the tissues changes with nutritional
state. In the fed, or postprandial, state there is a drive to store excess nutrients
and as part of this, adipose tissue LPL is up-regulated, probably by the action
of insulin released in response to carbohydrate in the meal. This diverts
chylomicron-fatty acids to adipose tissue rather than muscle, where LPL
is somewhat down-regulated by insulin (muscle LPL becomes particularly
active with physical training or during fasting, conditions when the muscle
has a greater need for fatty acids). Since adipose tissue is acquiring fatty acids
in the postprandial period, it makes sense that fat mobilization is suppressed
at this time, again by the action of insulin. Both the liver and skeletal muscle
take up fatty acids largely according to their availability. The rate of removal
of plasma NEFA is, under most conditions, fairly closely proportional to
their plasma concentration. The removal by the liver of remnant-particle
triacylglycerol-fatty acids is also probably determined by their delivery in
plasma. The abundance of the different sources of fatty acids in plasma varies
with nutritional state, with the highest supply of NEFA in the fasting state and
chylomicron-remnant triacylglycerol fatty acids becoming more prominent in
the fed state. As mentioned earlier, muscle LPL is up-regulated during fasting
although its activity does not seem to vary greatly during the normal day.
The secretion of VLDL-triacylglycerol by the liver is undoubtedly subject to
nutritional regulation. Insulin acutely suppresses hepatic VLDL-triacylglycerol
secretion. VLDL particles are secreted with a range of sizes, and it is especially
the secretion of the larger, more triacylglycerol-rich particles that is suppressed
by insulin. The mechanism by which insulin suppresses VLDL-triacylglycerol
secretion is 2-fold. First, it suppresses delivery of NEFA to the liver from
adipose tissue as described above. But there is a further, and more important,
effect on VLDL particle secretion from the hepatocytes. Whether insulin has this
effect in the normal daily feeding situation is not so clear: it is difficult to study in
vivo in humans because the non-steady state of feeding is not ideal for the tracer
experiments that have mostly been used to study regulation of VLDL secretion.
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Tissue-specific features of triacylglycerol and fatty acid metabolism
Adipose tissue
Adipose tissue triacylglycerol storage varies widely between individuals.
The ‘standard’ adult female has approx. 30% of body weight as fat and the
male approx. 20%, most of which is in adipocytes [1]. The adipocyte is a cell
highly specialized for fat storage, although we now also appreciate that it
has important protein secretory functions secreting hormones such as leptin,
adiponectin and a range of other so-called ‘adipokines’. The specific fatty acids
stored in human adipose tissue (saturated, monounsaturated and unsaturated)
reflect dietary intake. There is debate about the contribution of DNL (de novo
lipogenesis) to adipose tissue fat stores but, at the whole-body level, DNL only
operates significantly when carbohydrate intake exceeds energy requirements.
Isotopic data suggest that approx. 10% of adipose tissue triacylglycerol fatty
acids arise from DNL [2]. Most of the triacylglycerol-fatty acids in human
adipose tissue therefore arise from plasma, mainly from the triacylglycerol
fraction of plasma lipids with a small contribution of direct uptake from
NEFAs [3].
The control of fatty acid uptake by adipocytes is not well understood.
Up-regulation of adipose tissue LPL activity in the fed state is brought about
by multiple mechanisms (Figure 2). LPL is synthesized within adipocytes
and exported to its site of action, the luminal side of the capillary endothelial
cells. Of the large pool of LPL in adipose tissue, only a portion becomes active
endothelial-bound LPL. An important mechanism for up-regulation in the
fed state is the diversion of LPL away from the degradative, into the secretory
pathway [4]. LPL acts on circulating lipoprotein-triacylglycerol and generates
fatty acids in the local area close to the endothelium. Chylomicrons are the preferred substrate for adipose tissue LPL. Chylomicron-triacylglycerol is hydrolysed considerably more rapidly than VLDL-triacylglycerol. Nevertheless,
both substrates may contribute fatty acid under appropriate circumstances.
The fatty acids released by LPL migrate across the endothelial wall and into
the underlying adipocytes. The mechanisms involved in this process are not
clear, although there is increasing evidence for the involvement of specific fatty
acids transporters including FAT (fatty acid translocase, also known as CD36)
in facilitating fatty acid movement across the adipocyte cell membrane. There
is also non-regulated diffusion through the cell membrane. Within the adipocyte, fatty acids are ‘activated’ by esterification to CoA in an ATP-requiring
process that seems to be intimately linked to fatty acid transport into the cell,
at least when mediated by a member of the FATP (fatty acid transport protein)
family (FATP1) [5].
Adipose tissue LPL generates an excess of fatty acids. The adipocytes then
take up a proportion, depending upon nutritional state. There is always an
‘overspill’ of fatty acids into the plasma. Insulin increases the proportion of
LPL-derived fatty acids taken up by the tissue but even in the postprandial state
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Catecholamines
Insulin
Natriuretic peptides Tumour necrosis factor-α
Adrenergic
receptor
Insulin
receptor
Natriuretic
peptide
receptor A
TNF-α
receptor 1
Cyclic GMP
MAP kinases
Phosphodiesterase 3B
Cyclic AMP
5′AMP
Protein
kinase A
Protein
kinase G
perilipin
?
Triacylglycerol
lipase
Triacylglycerol
Hormone-sensitive
lipase
Monoacylglycerol
lipase
Diacylglycerol
Monoacylglycerol
Fatty acid
Fatty acid
Glycerol
Fatty acid
Figure 3. Regulation of lipolysis in white adipose tissue
For more detail please refer to the text.
typically approx. 50% may be released as NEFA, clearly shown by enrichment
of the plasma NEFA pool with dietary fatty acids soon after a meal. These
fatty acids are then available for uptake by other tissues including muscle and
liver. The control of fatty acid uptake into the adipocyte may involve recruitment of the fatty acid transporters CD36 and FATP1 to the cell surface upon
insulin stimulation [5]. It also involves stimulation of fatty acid esterification
to make new triacylglycerol within the adipocyte. Regulation of the enzymes
of triacylglycerol synthesis is still not understood in detail although the pathway as a whole is certainly stimulated by insulin [6]. In addition, insulin may
stimulate glucose uptake and glycolysis, which supplies the glycerol 3-phosphate necessary for fatty acid esterification. The ability of fat cells to take up
acylglycerols directly is very limited.
Adipose tissue fat mobilization has been studied in detail [7] and is summarized in Figure 3. Until recently it was considered that the only major
regulatory point was the enzyme HSL. HSL has triacylglycerol-lipase activity
directed at two fatty acids on a triacylglycerol molecule, but is not active against
monoacylglycerols, whereas it is especially active against diacylglycerols.
There is a separate, and unrelated, monoacylglycerol lipase that is consitutively
expressed in adipose tissue and catalyses the last step in triacylglycerol hydrolysis. HSL is regulated in the long-term by transcriptional control: HSL mRNA
abundance increases during starvation, and is decreased in obesity. But the most
important aspect of HSL on a day-to-day basis is its regulation by reversible
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phosphorylation [7]. HSL has three phosphorylation sites that are substrates for
PKA (protein kinase A), phosphorylation of which greatly increases its activity.
Adipocyte fat mobilization is brought about by signals that increase the cAMP
concentration. Typically, catecholamines would act by binding to ␤-adrenoceptors in the cell membrane, linked by G-proteins to adenylate cyclase.
Insulin suppresses HSL activity by activation of a particular phosphodiesterase, PDE3B, which reduces the cAMP concentration. These effects are rapid
and potent. However, the increased rate of lipolysis seen upon stimulation of
fat cells with ␤-adrenergic agents is far greater than would be predicted from
the increase in activity of HSL in vitro. It is now clear that phosphorylation of
HSL is accompanied by phosphorylation of a protein, perilipin, that coats the
adipocyte fat droplet. This dual phosphorylation allows HSL to translocate
from the cytosol to the surface of the fat droplet to achieve lipolysis. Under
non-stimulated conditions, HSL is excluded from the fat droplet surface.
A further system of human-specific regulation has been described, involving the circulating peptides known as ANF (atrial natriuretic factor) and BNF
(brain natriuretic factor) [8]. These factors are released during stress states
including exercise and bind to specific cell-surface receptors that are coupled to synthesis of cGMP and activation of cyclic GMP-dependent protein
kinase. Finally, there is an autocrine regulation of lipolysis by tumour necrosis
factor-␣. This cytokine acts on perilipin through MAPK (mitogen-activated
protein kinase) signalling pathways.
The primary role of HSL in lipolysis has been challenged by findings in
mice deficient in HSL. These mice have relatively normal adipose tissue mass,
and lipolysis is preserved, although ␤-adrenergic responsiveness is almost or
completely lost, depending upon the model. This has led to a search for other
lipases in the adipocyte. One candidate, ATGL (adipose triglyceride lipase), has
marked specificity for triacylglycerols compared with di- or mono-acylglycerols,
and may be of great importance for lipolysis in rodents. It now seems likely that
ATGL is responsible for ‘basal’ lipolysis in human adipose tissue, with HSL
responsible for the majority of catecholamine- or ANF-stimulated lipolysis [9].
Liver
The normal triacylglycerol content of the liver is around 1–10% by weight
[1] but can expand considerably in the condition generally called ‘fatty liver’.
The liver receives plasma fatty acids as NEFA, from hydrolysis of circulating
triacylglycerol and from the uptake of remnant lipoprotein particles. Uptake
of NEFA appears to be similar to the process described in adipose tissue:
FAT/CD36, FATP2 and FATP5 are expressed in liver. The adult liver
does not express LPL, but a related enzyme, HL (hepatic lipase) which acts
preferentially upon smaller particles than LPL. It will remove triacylglycerol
from small VLDL particles and from particles on their way to becoming LDL
(low-density lipoprotein) and also from HDL (high-density lipoprotein)
particles. It also has a major role in ‘docking’ with particles that are being
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internalized bound to specific receptors expressed in the liver. The liver is
the main organ for uptake of the remnant particles formed from the action
of LPL on triacylglycerol-rich chylomicrons and VLDL. The chylomicron
remnant particle is thought to contain typically about one-third of its original
triacylglycerol content. This is still appreciable (a typical chylomicron particle
contains approx. 106 molecules of triacylglycerol), and would lead to delivery
to the liver of around 20–30 g/day of dietary triacylglycerol.
In addition, the liver has the enzymatic capacity for DNL, the synthesis of
fatty acids from glucose and other non-lipid precursors. DNL makes a small
contribution in people on a typical, western relatively high-fat diet, but it becomes
activated during a high-energy diet rich in carbohydrate [10]. The pathway
involves export of acetyl-CoA formed in the mitochondrion (in the case of
glucose as the precursor, by pyruvate dehydrogenase) to the cytosol (in the form
of citrate), conversion of acetyl-CoA to malonyl-CoA by acetyl-CoA carboxylase
and then sequential addition of two carbon units by fatty acid synthase.
Fatty acids taken up into the hepatocytes are activated by esterification to
CoA. The long-chain acyl-CoA may then enter the mitochondria for oxidation
or be a substrate for glycerolipid synthesis (triacylglycerol and phospholipids).
This is a key regulatory point, determined largely by regulation of entry into the
mitochondria, via the enzyme CPT 1 (carnitine palmitoyltransferase 1, the ‘overt’
isoform, expressed in the outer mitochondrial enzyme). This enzyme is allosterically regulated by the cytosolic malonyl-CoA concentration. Malonyl-CoA
(an intermediate in fatty acid synthesis) is a very potent inhibitor of CPT 1. Its
concentration is high when insulin levels are high and glucose is readily available,
conditions when fatty acid synthesis should be favoured over fatty acid oxidation.
In the fasting state, when insulin levels are low, fatty acid oxidation is therefore
favoured. There is a further branch point in fatty acid oxidation when acetyl-CoA
may either enter the tricarboxylic acid cycle or may be directed into ketone body
synthesis. Less is known about regulation of this ‘metabolic branch point’. A
suggested regulation of ketone body synthesis is by reversible succinylation of
the mitochondrial 3-hydroxy-3-methylglutaryl-CoA synthase [11].
The cytosolic triacylglycerol pool within the hepatocyte is the precursor
for VLDL-triacylglycerol. Lipolysis of this cytosolic pool is necessary to generate fatty acids that are then esterified to make new triacylglycerol within the
endoplasmic reticulum, where VLDL particles are assembled. The identity of
the lipase responsible, and its regulation, are not clear. VLDL-triacylglycerol
secretion is acutely inhibited by insulin, although hepatic fatty acids are preferentially stored as triacylglycerol when insulin levels are high. This implies that
the hepatocyte triacylglycerol store fluctuates in size, increasing after meals
and then discharging during fasting as VLDL are secreted. This has recently
been visualized directly in healthy humans where approx. 10% of the fat from
a meal was stored in the liver in the few hours following feeding [12].
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Muscle
The total triacylglycerol store in a healthy human liver is approx. 100 g (or
less). The typical triacylglycerol content in whole-body skeletal muscle might
be approx. 300 g (estimated making various assumptions, see [13]).
There are two major triacylglycerol stores in muscle: in adipocytes interleaved with the muscle fibres (seen, for instance, as ‘marbling’ in a cut of meat)
and triacylglycerol stored within the muscle fibre itself (‘intramyocellular triacylglycerol’). Here we will discuss only the latter. It is still not clear whether
there is any direct metabolic relationship between adipocytes and skeletal
muscle fibres even when they are located in close proximity.
Intramyocellular triacylglycerol is present as lipid droplets, often in close
contact with mitochondria. Some of the enzymatic machinery for fatty acid and
triacylglycerol metabolism in the muscle fibre is similar to that in the adipocyte.
Fatty acids can be taken up from the plasma, either from the albumin-bound
NEFA pool or, as in adipose tissue, from the pool of fatty acids generated by
the action of LPL on circulating triacylglycerol. In either case it seems that the
fatty acids enter the cells at least in part by facilitated diffusion, as both FAT/
CD36 and FATP1 are expressed in skeletal muscle. Muscle has the enzymatic
machinery to synthesize malonyl-CoA from glucose utilization, the first stage of
the biosynthesis of fatty acids. However, expression of fatty acid synthase (which
makes fatty acids from malonyl-CoA) is very small or non-existent. Therefore
the triacylglycerol present in muscle can be assumed to arise entirely from plasma
fatty acids (NEFA or triacylglycerol). The role of malonyl-CoA appears to be in
the regulation of fatty acid oxidation, as described above for the liver.
The intramyocellular triacylglycerol is hydrolysed to make fatty acids
available for oxidation. HSL is likely to be the key lipase controlling this process, although there is no direct evidence for this other than the fact that HSL
is expressed in muscle and becomes activated on muscle contraction [14]. This
is reinforced by the accumulation of large lipid droplets in skeletal muscle of
HSL-deficient mice [15]. Fatty acids made available from muscle triacylglycerol lipolysis would be a substrate for oxidation after activation and entry
into the mitochondria. Muscle lipolysis is less sensitive to hormone regulation than adipocyte lipolysis, at least in humans. Insulin is not effective but
catecholamines stimulate muscle lipolysis [14,16].
It is not clear whether, as in the adipocyte, fatty acids may be released
from the cell into the plasma. Certainly this does not happen in a net sense:
muscle always extracts NEFA from plasma, never adds them. Some measurements suggest that fatty acids are released from muscle beds, in larger amounts
than could be accounted for by the adipocytes likely to be present [17].
Nevertheless, it is clear that adipose tissue is the only tissue that contributes
NEFA to the plasma in a net sense.
The processes of fatty acid uptake by muscle, and of mobilization of
muscle triacylglycerol stores, are covered in detail in Chapter 4 and will not be
covered here in further detail.
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The heart muscle (myocardium) is also involved in fatty acid metabolism;
in fact, as a continuously contracting oxidative muscle, it has a very high
demand for fatty acids. These fatty acids are acquired from plasma both as
NEFA and through the LPL-route. Many features of myocardial fatty acid
metabolism are similar to skeletal muscle, although the major isoform of the
FATP family expressed in the heart is FATP6, which co-localizes on the plasma
membrane with CD36.
Metabolic interactions during exercise
Fat is an important fuel for skeletal muscle during exercise. Its importance is
greater at lower intensities of exercise and as the duration of exercise increases.
In any exercise lasting more than approximately 2 h, fat becomes the major
fuel (once muscle glycogen stores are exhausted). Most of the fatty acids
oxidized in muscle during exercise come from the plasma NEFA pool. Hence
coordination between adipose tissue (activation of lipolysis) and muscle (fatty
acid utilization) is crucial. The contribution of VLDL-triacylglycerol fatty
acids during exercise is small, but in the fed state chylomicron-triacylglycerol
fatty acids might make a greater contribution [18]. Regarding the role of the
intramuscular triacylglycerol store, early studies involving muscle biopsies
before and after exercise were generally uninformative. New approaches
have increased our understanding of muscle triacylglycerol. Isotope infusion
studies show that muscle triacylglycerol utilization makes a substantial
contribution to fatty acid oxidation at moderate intensity of exercise (approx.
55–65% VO2max), almost equal to the contribution from plasma NEFA,
but less at lesser or greater intensity [19,20]. The measurement of IMCL
(intra-myocellular lipid) by magnetic resonance spectroscopy also shows
muscle triacylglycerol utilization during exercise, at rates comparable with
those observed with tracer techniques [21,22].
The activation of adipose tissue lipolysis during exercise is largely
␤-adrenergic, responding presumably to circulating adrenaline or perhaps
activation of the sympathetic outflow to adipose tissue (studies in people
with spinal cord injuries suggest mainly the former [23]). The technique of
microdialysis of adipose tissue involves a small probe of semi-permeable membrane, which is used both to introduce agents and to sample the interstitial
glycerol concentration (as a marker of lipolysis). Introduction of propanolol
(a non-specific ␤-adrenergic blocker) largely, but not completely, inhibits the
stimulation of lipolysis by exercise [24]. The remaining, non-␤-adrenergic
component, may represent activation by the ANF pathway [8].
If activation of adipose tissue lipolysis relies on signals that are part of the
exercise response, how is it that the rate of fatty acid delivery so closely matches
the rate of utilization by skeletal muscle? This would seem to require some
direct signal from the muscle to the adipose tissue. If someone exercises whilst
lipolysis is suppressed, e.g. using the drug acipimox which acts on the so-called
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nicotinic acid receptor on adipocytes and potently blocks NEFA release, then
the muscles use more IMCL and also more glucose. That implies that NEFA
delivery from adipose tissue is a ‘preferred’ fuel and the muscles will use other
fuels to make up any lack of NEFA. An alternative view is that adipose tissue
lipolysis normally operates at a greater rate than is necessary for fatty acid
oxidation in other tissues. The ‘excess’ fatty acids are re-esterified, for instance
in the liver, and returned to adipose tissue in VLDL. Then, at the onset of
exercise, a greater proportion of the NEFA delivered from adipose tissue will
be oxidized and a correspondingly lower portion re-esterified [25]. However,
this view is not supported by all studies [17].
Tissue triacylglycerol metabolism in health and disease
The systems described above function efficiently in health to maintain
appropriate delivery of fatty acids when they are needed. Dysfunction of
these systems may underlie some specific disorders of lipid metabolism. For
instance, in the relatively common condition known as familial combined
hyperlipidaemia, there is evidence that the primary defect is an over-production
of small VLDL particles from the liver. This may be combined with a low rate
of lipolysis in adipose tissue [26].
We will discuss here one very common condition which may arise from a
dysfunction of these systems: the ‘metabolic syndrome’ or ‘insulin resistance
syndrome’. This is a collection of inter-connected adverse metabolic changes
including insulin resistance (with some degree of glucose intolerance), elevated
blood triacylglycerol concentrations and low concentrations of the ‘protective’
HDL-cholesterol. This condition is usually associated with central (abdominal)
obesity. It has become clear in recent years that insulin resistance is closely
associated with increased concentrations of triacylglycerol in liver and skeletal
muscle. An increase in triacylglycerol deposition in the pancreatic ␤-cell may
lead, in time, to impairment of insulin secretion, therefore explaining why this
syndrome is an important precursor to development of type 2 diabetes. This
has been termed ‘ectopic fat deposition’, i.e. deposition of triacylglycerol in tissues other than adipose tissue. It may arise from a dysfunction of the adipose
tissue, leading to exposure of other tissues to excess flux of fatty acids (both
NEFA and triacylglycerol-rich remnant particles) [27]. One possible mechanism is that abdominal fat, including the visceral fat within the abdominal
cavity, is less efficient than is lower-body adipose tissue at ‘sequestering’ excess
dietary fat and thereby protecting other tissues. There is evidence for this view
from tracer studies showing a high rate of turnover of triacylglycerol stores
in abdominal, especially intra-abdominal adipose tissue [28] and indirectly
from the strong protective effect of lower-body adipose tissue against insulin
resistance, dyslipidaemia and hypertension [29].
Why should an increased triacylglycerol concentration in liver and skeletal
muscle be associated with insulin resistance? The difficulty is in dissociating
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cause and effect. It is possible that the persistent high insulin concentrations
that characterize insulin resistance lead to altered partitioning of fatty acids,
with more going towards esterification at the expense of oxidation. In that
view, ectopic fat deposition would be a consequence of insulin resistance rather
than a cause. But there are also strong suggestions of a causal relationship.
There are many demonstrations, especially in skeletal muscle (which is more
accessible to study than the liver), that elevated fatty acid availability reduces
the efficiency of insulin signalling (e.g. to stimulate glucose metabolism).
The underlying mechanisms are discussed in Chapter 4 of this volume. The
potential mechanisms by which regular exercise and training prevent insulin
resistance despite large intramyocellular triacylglycerol stores in muscle are
discussed in several of the essays in this volume.
It is also clear that treatments that reduce ‘ectopic’ fat content improve
insulin resistance. For instance, the TZD (thiazolidinedione) insulin-sensitizing
drugs that are now in clinical use for the treatment of type 2 diabetes improve
insulin sensitivity and reduce hepatic fat content [30]. The interesting point is
that the TZDs are agonists to the nuclear receptor/transcription factor PPAR␥
(peroxisome-proliferator-activated receptor ␥), which is most highly expressed
in adipose tissue and not expressed in liver. This suggests that TZDs reduce
liver fat and improve hepatic insulin sensitivity by effects on adipose tissue,
possibly increasing the capacity of adipose tissue to ‘entrap’ fatty acids and
thus protect the liver.
There are further links between the tissues that are the subject of this
chapter. It is now recognized that adipose tissue secretes a large number of
peptides and other factors, some of which may function as hormones and
affect metabolism in other tissues. One of these is a protein called adiponectin. It is abundant in the circulation (typical plasma concentration 10 mg/l).
Adiponectin is most highly secreted by small adipocytes. As adipocytes
increase in size, adiponectin expression and secretion are so highly downregulated (the mechanism is unknown) that plasma concentrations are actually
lower in obese than in lean people. There are strong relationships between
high adiponectin concentrations and a ‘healthy’ metabolic profile, or conversely between low adiponectin and features of the metabolic syndrome. At
least some of these effects may be mediated by adiponectin-induced reduction
in liver fat content. The molecular mechanism may involve activation of the
AMP-activated protein kinase [31]. It is also noteworthy that a strong effect
of TZD administration is to raise adiponectin expression, secretion and plasma
concentration, and that the reduction in liver fat content observed with TZD
treatment is closely related to the increase in adiponectin concentration [30].
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Essays in Biochemistry volume 42 2006
Summary
•
•
•
•
•
Fat is the most important stored fuel in mammals.
Three tissues play major roles in fatty acid and triacylglycerol metabolism: adipose tissue, skeletal muscle and liver.
Each of these tissues has a store of triacylglycerol that can be hydrolysed (mobilized) in a regulated way to release fatty acids.
There is clearly cooperation amongst the tissues, so that, for instance,
adipose tissue fat mobilization increases to meet the demands of skeletal muscle during exercise.
When triacylglycerol accumulates excessively in skeletal muscle and
liver, sometimes called ectopic fat deposition, then the condition of
insulin resistance arises. This may reflect a lack of exercise and an excess
of fat intake.
The authors collaborate as part of the project ‘Hepatic and adipose tissue and
functions in the metabolic syndrome’ (HEPADIP, http://www.hepadip.org/), which
is supported by the European Commission as an Integrated Project under the 6th
Framework Programme (Contract LSHM-CT-2005-018734).
References
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
Snyder, W.S. (1975) Report of the task force on reference man, Pergamon Press for the International
Commission on Radiological Protection, Oxford
Strawford, A., Antelo, F., Christiansen, M. & Hellerstein, M.K. (2004) Adipose tissue triglyceride
turnover, de novo lipogenesis, and cell proliferation in humans measured with 2H2O. Am. J. Physiol.
Endocrinol. Metab. 286, E577–E588
Coppack, S.W., Persson, M., Judd, R.L. & Miles, J.M. (1999) Glycerol and nonesterified fatty acid
metabolism in human muscle and adipose tissue in vivo. Am. J. Physiol. 276, E233–E240
Bergö, M., Olivecrona, G. & Olivecrona, T. (1996) Forms of lipoprotein lipase in rat tissues: in
adipose tissue the proportion of inactive lipase increases on fasting. Biochem. J. 313, 893–898
Stahl, A. (2004) A current review of fatty acid transport proteins (SLC27). Pflügers Arch. 447,
722–727
Coleman, R.A. & Lee, D.P. (2004) Enzymes of triacylglycerol synthesis and their regulation. Prog.
Lipid Res. 43, 134 –176
Holm, C. (2003) Molecular mechanisms regulating hormone-sensitive lipase and lipolysis. Biochem.
Soc. Trans. 31, 1120 –1124
Lafontan, M., Moro, C., Sengenes, C., Galitzky, J., Crampes, F. & Berlan, M. (2005) An unsuspected
metabolic role for atrial natriuretic peptides: the control of lipolysis, lipid mobilization, and systemic nonesterified fatty acids levels in humans. Arterioscler. Thromb. Vasc. Biol. 25, 2032 – 2042
Langin, D., Dicker, A., Tavernier, G., Hoffstedt, J., Mairal, A., Rydén, M., Arner, E., Sicard, A.,
Jenkins, C.M., Viguerie, N. et al. (2005) Adipocyte lipases and defect of lipolysis in human obesity.
Diabetes 54, 3190 –3197
Hellerstein, M.K., Schwarz, J.-M. & Neese, R.A. (1996) Regulation of hepatic de novo lipogenesis
in humans. Annu. Rev. Nutr. 16, 523 – 557
Hegardt, F.G. (1999) Mitochondrial 3-hydroxy-3-methylglutaryl-CoA synthase: a control enzyme
in ketogenesis. Biochem. J. 338, 569 – 582
© 2006 The Authors
Ch-07_essbiochem_frayn.indd
102
11/4/06
5:13:25 PM
K.N. Frayn, P. Arner & H. Yki-Järvinen
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
29.
30.
31.
32.
103
Ravikumar, B., Carey, P.E., Snaar, J.E.M., Deelchand, D.K., Cook, D.B., Neely, R.D.G., English, P.T.,
Firbank, M.J., Morris, P.G. & Taylor, R. (2005) Real-time assessment of postprandial fat storage
in liver and skeletal muscle in health and type 2 diabetes. Am. J. Physiol. Endocrinol. Metab. 288,
E789–E797
Frayn, K.N. & Blaak, E.E. (2005) Metabolic fuels and obesity: carbohydrate and lipid metabolism
in skeletal muscle and adipose tissue, in: Clinical Obesity in Adults and Children (Kopelman, P.,
Caterson, I., Dietz, W., eds), pp. 102 –122, Blackwell Publishing Ltd, Oxford
Donsmark, M., Langfort, J., Holm, C., Ploug, T. & Galbo, H. (2005) Hormone-sensitive lipase as
mediator of lipolysis in contracting skeletal muscle. Exerc. Sport. Sci. Rev. 33, 127–133
Hansson, O., Donsmark, M., Ling, C., Nevsten, P., Danfelter, M., Andersen, J.L., Galbo, H. &
Holm, C. (2005) Transcriptome and proteome analysis of soleus muscle of hormone-sensitive
lipase-null mice. J. Lipid Res. 46, 2614 – 2623
Moberg, E., Sjöberg, S., Hagström-Toft, E. & Bolinder, J. (2002) No apparent suppression by
insulin of in vivo skeletal muscle lipolysis in nonobese women. Am. J. Physiol. Endocrinol. Metab.
283, E295–E301
van Hall, G., Sacchetti, M., Radegran, G. & Saltin, B. (2002) Human skeletal muscle fatty acid and
glycerol metabolism during rest, exercise and recovery. J. Physiol. 543, 1047–1058
Henriksson, J. (1995) Muscle fuel selection: effect of exercise and training. Proc. Nutr. Soc. 54,
125–138
Romijn, J.A., Coyle, E.F., Sidossis, L.S., Gastaldelli, A., Horowitz, J.F., Endert, E. & Wolfe, R.R.
(1993) Regulation of endogenous fat and carbohydrate metabolism in relation to exercise
intensity and duration. Am. J. Physiol. 265, E380 –E391
van Loon, L.J.C., Greenhaff, P.L., Constantin-Teodosiu, D., Saris, W.H.M. & Wagenmakers, A.J.M.
(2001) The effects of increasing exercise intensity on muscle fuel utilisation in humans. J. Physiol.
536, 295 –304
Watt, M.J., Heigenhauser, G.J. & Spriet, L.L. (2002) Intramuscular triacylglycerol utilization in
human skeletal muscle during exercise: is there a controversy? J. Appl. Physiol. 93, 1185–1195
van Baak, M.A., Mooij, J.M.V. & Wijnen, J.A.G. (1993) Effect of increased plasma non-esterified
fatty acid concentrations on endurance performance during ␤-adrenoceptor blockade. Int. J.
Sports Med. 14, 2– 8
Stallknecht, B., Lorentsen, J., Enevoldsen, L.H., Bülow, J., Biering-Sørensen, F., Galbo, H. & Kjaer,
M. (2001) Role of the sympathoadrenergic system in adipose tissue metabolism during exercise in
humans. J. Physiol. 536, 283 – 294
Arner, P., Kriegholm, E., Engfeldt, P. & Bolinder, J. (1990) Adrenergic regulation of lipolysis in situ
at rest and during exercise. J. Clin. Invest. 85, 893 – 898
Wolfe, R.R., Klein, S., Carraro, F. & Weber, J.-M. (1990) Role of triglyceride-fatty acid cycle in
controlling fat metabolism in humans during and after exercise. Am. J. Physiol. 258, E382 – E389
Reynisdottir, S., Eriksson, M., Angelin, B. & Arner, P. (1995) Impaired activation of adipocyte
lipolysis in familial combined hyperlipidemia. J. Clin. Invest. 95, 2161– 2169
Frayn, K.N. (2002) Adipose tissue as a buffer for daily lipid flux. Diabetologia 45, 1201–1210
Jensen, M.D., Sarr, M.G., Dumesic, D.A., Southorn, P.A. & Levine, J.A. (2003) Regional uptake of
meal fatty acids in humans. Am. J. Physiol. Endocrinol. Metab. 285, E1282 – E1288
Lemieux, I. (2004) Energy partitioning in gluteal-femoral fat: does the metabolic fate of
triglycerides affect coronary heart disease risk? Arterioscler. Thromb. Vasc. Biol. 24, 795 – 797
Tiikkainen, M., Häkkinen, A.M., Korsheninnikova, E., Nyman, T., Mäkimattila, S. & Yki-Järvinen,
H. (2004) Effects of rosiglitazone and metformin on liver fat content, hepatic insulin resistance,
insulin clearance, and gene expression in adipose tissue in patients with type 2 diabetes. Diabetes
53, 2169–2176
Yamauchi, T., Kamon, J., Minokoshi, Y., Ito, Y., Waki, H., Uchida, S., Yamashita, S., Noda, M., Kita,
S., Ueki, K. et al. (2002) Adiponectin stimulates glucose utilization and fatty-acid oxidation by
activating AMP-activated protein kinase. Nat. Med. 8, 1288 –1295
Frayn, K.N. (2003) Metabolic regulation: A human perspective. 2nd edn, Blackwell Publishing, Oxford
© 2006 The Authors
Ch-07_essbiochem_frayn.indd
103
11/4/06
5:13:25 PM